Laboratory Techniques: Slide Preparation and

Transcription

Glime, J. M. and Wagner, D. H. 2013. Laboratory Techniques: Slide Preparation and Stains. Chapt. 2-2. In: Glime, J. M. Bryophyte
Ecology. Volume 3. Methods. Ebook sponsored by Michigan Technological University and the International Association of Bryologists.
Ebook last updated 11 September 2013 and available at <www.bryoecol.mtu.edu>.
2-2-1
CHAPTER 2-2
LABORATORY TECHNIQUES:
SLIDE PREPARATION AND STAINS
TABLE OF CONTENTS
Preparing the Specimen....................................................................................................................................... 2-2-3
Cleaning Bryophytes.................................................................................................................................... 2-2-3
Washing Machine ................................................................................................................................. 2-2-3
Embroidery Hoop ................................................................................................................................. 2-2-3
Wash Bottle........................................................................................................................................... 2-2-3
HCl........................................................................................................................................................ 2-2-4
Aquatic Bryophytes .............................................................................................................................. 2-2-4
Sorting the Plants ......................................................................................................................................... 2-2-4
Wetting Agents ............................................................................................................................................ 2-2-4
Soap ...................................................................................................................................................... 2-2-5
Agral 600 .............................................................................................................................................. 2-2-5
Clearing Leaves ........................................................................................................................................... 2-2-5
Lactic Acid............................................................................................................................................ 2-2-5
KOH or NaOH ...................................................................................................................................... 2-2-6
Chloral Hydrate..................................................................................................................................... 2-2-6
Dehydration.................................................................................................................................................. 2-2-6
Stains............................................................................................................................................................ 2-2-6
Staining Stems ...................................................................................................................................... 2-2-7
Triple Stains................................................................................................................................... 2-2-7
Kawai Stem Staining Techniques .................................................................................................. 2-2-7
Acid Fuchsin................................................................................................................................ 2-2-14
Aniline Blue................................................................................................................................. 2-2-14
Congo Red ................................................................................................................................... 2-2-14
Eosin ............................................................................................................................................ 2-2-14
Fast Green.................................................................................................................................... 2-2-14
Fuchsin......................................................................................................................................... 2-2-14
Gentian Violet (=Crystal Violet) ................................................................................................. 2-2-14
Janus Green.................................................................................................................................. 2-2-14
Methyl Green ............................................................................................................................... 2-2-14
Leaves ................................................................................................................................................. 2-2-15
I2KI .............................................................................................................................................. 2-2-15
KOH or NaOH............................................................................................................................. 2-2-15
Methylene Blue............................................................................................................................ 2-2-15
Safranin O / Fast Green ............................................................................................................... 2-2-15
Sphagnum Stains................................................................................................................................. 2-2-15
Methylene Blue............................................................................................................................ 2-2-15
Crystal Violet/Gentian Violet ...................................................................................................... 2-2-16
Toluidine Blue O ......................................................................................................................... 2-2-16
Reproductive Structures...................................................................................................................... 2-2-16
Iron Haematoxylon / Fast Green.................................................................................................. 2-2-16
Bulbils and Spores .............................................................................................................................. 2-2-16
Fluorescence and Fluorescent Dyes............................................................................................. 2-2-16
Staining Liverwort Capsules............................................................................................................... 2-2-17
pH Testing.................................................................................................................................................. 2-2-18
Weak Alkali ............................................................................................................................................... 2-2-18
Cleaning Up Stains..................................................................................................................................... 2-2-19
2-2-2
Chapter 2-2: Lab Techniques: Slide Preparation and Stains
Leaf Removal and Making Slides..................................................................................................................... 2-2-19
Sectioning ......................................................................................................................................................... 2-2-19
Razor Blades.............................................................................................................................................. 2-2-19
Cutting Techniques.................................................................................................................................... 2-2-19
Wax Mounts ....................................................................................................................................... 2-2-20
Cutting Block ..................................................................................................................................... 2-2-19
Pith Sandwich Cutting Tool ............................................................................................................... 2-2-20
Chopping Method............................................................................................................................... 2-2-21
Roll and Chop..................................................................................................................................... 2-2-21
Modified Roll and Chop..................................................................................................................... 2-2-22
Dissecting Microscope Hand Sections ............................................................................................... 2-2-22
Double Slide Sectioning Technique ................................................................................................... 2-2-22
Cryostat Sections................................................................................................................................ 2-2-24
Stems and Small Leaves ............................................................................................................................ 2-2-24
Techniques for Special Structures .................................................................................................................... 2-2-25
Clearing Spores ......................................................................................................................................... 2-2-25
SEM........................................................................................................................................................... 2-2-25
Vacuoles .................................................................................................................................................... 2-2-26
Liverworts.................................................................................................................................................. 2-2-26
Peristome Teeth ......................................................................................................................................... 2-2-26
Summary........................................................................................................................................................... 2-2-27
Acknowledgments ............................................................................................................................................ 2-2-27
Literature Cited................................................................................................................................................. 2-2-27
Glime, J. M. and Wagner, D. H. 2013. Laboratory Techniques: Slide Preparation and Stains. Chapt. 2-2. In: Glime, J. M. Bryophyte
Ecology. Volume 3. Methods. Ebook sponsored by Michigan Technological University and the International Association of Bryologists.
Ebook last updated 11 September 2013 and available at <www.bryoecol.mtu.edu>.
2-2-3
CHAPTER 2-2
LAB TECHNIQUES:
SLIDE PREPARATION AND STAINS
Figure 1. Polytrichum juniperinum leaf cross section using a cryostat and displaying natural colors. Photo by John Hribljan.
Preparing the Specimen
Fresh specimens are the most fun to work with. They
are bright green and require little or no hydration before
placing them in a drop of water on a slide. Chloroplasts
migrate in cyclosis. And tiny invertebrates crawl about to
entertain and distract you. But most often we don't have
the pleasure to observe fresh material under the microscope.
Instead, we have dry, often brittle, specimens collected in
great numbers in a day-long or even months-long collecting
trip. But don't dismay – the bryophytes will still freshen up
to make good slides.
Cleaning Bryophytes
Washing Machine (Jewett 1913)
Jewett (1913) suggests a small "washing machine."
The bryophytes are placed on a fine screen – we assume
that cloth window screening would work – and sprayed
with a nozzle to clean them.
Embroidery Hoop (Mayfield et al. 1983)
Mayfield et al. (1983) suggested a similar cleaning
procedure using a net, but they suggested placing the
netting (mosquito or bridal veil netting) tightly in an
embroidery hoop. This is particularly useful for thallose
liverworts. They should be collected with ~3 mm substrate
to protect rhizoids and scales. The liverworts are placed on
the hoop netting with a second net placed over them. They
are then washed with a stream of water. This may take
some practice because too much water will damage the
plants whereas a weak stream will not succeed in removing
the soil and debris. Mayfield and coworkers suggest that a
suitable stream of water can be achieved by attaching an
eyedropper to pliable tubing. If the tubing is connected to a
tapered laboratory water faucet, water flow can be
adequately controlled. Specimens can then be pressed
suitably in a telephone book, using folded waxed paper to
hold the specimens between the pages of the book. Dried
specimens are affixed to a 2x5" (5x12.5 cm) card with
water-soluble glue. Specimens can be rehydrated when
needed with boiling water.
Contemporary workers
discourage pressing or gluing specimens.
Wash Bottle (Wagner 2011)
Wagner (2011) suggests having a small wash bottle
(125 ml) for rinsing the bryophytes and cleaning slides and
coverslips for reuse (Figure 2). The water can also be used
to wash away the wetting agent. The same ability of a
wetting agent (see below) to reduce trapped bubbles also
causes the water drop on your slide to lose its cohesion and
adhesion, causing the water drop to run all over the place,
so start with a small drop.
2-2-4
Figure 3. Experiment on cleaning Fontinalis sp. with
household hydrogen peroxide at various concentrations. Note the
bleaching at 70 and 80%. Photo by Janice Glime.
Sorting the Plants
Figure 2. Water bottle and ceramic washing cup. David
Wagner says he likes "to use a pretty, wood-fired stoneware cup,
much more pleasing to the eye than the usual beaker." Photo by
David Wagner.
HCl (Zander 1993)
Zander (1993) suggests using dilute HCl to clean away
limy incrustations. It can also indicate whether the
collection was made from a calcareous habitat because, if it
is calcareous and bits are present with the sample, it will
produce bubbles.
Aquatic Bryophytes (Landry 1973)
Aquatic bryophytes can be particularly challenging.
They typically are covered with epiphytes, some of which
(e.g., the diatom Cocconeis) embed themselves into the
surfaces of the leaves. Landry (1973) experimented with
various cleaning techniques on Fontinalis. He found that
household bleach, diluted to 0.5%, causes no chlorophyll
bleaching, but at 0.10% bleaching appeared in 15 seconds.
Solutions diluted to 0.25% caused bleaching in 5 seconds.
Bleaching in these cases occurred in the lower (older)
leaves and may have been tied to senescence.
Unfortunately, these methods did not appear to remove the
epiphytes. Ultimately, 5 minutes cleaning with 3%
peroxide (H2O2) and agitation seemed to remove
approximately 85-90% of the epiphytes (Figure 3).
Tumbling the moss at 30 revolutions per minute still only
removed only about 85% of the epiphytes. Swirling
improved the removal. Higher concentrations of peroxide
and/or longer time periods caused bleaching of the
chlorophyll. Based on the improved success with agitation,
Landry and Glime (unpublished) tried an ultrasound bath.
This improved the removal of the epiphytes, but the
internal cell structure of the moss was disrupted.
A classic mistake in identifying bryophytes is looking
at the sporophyte of one species and the leaves of another.
Sporophytes often originate deep in the clump and may
actually belong to a species that achieved sufficient
dominance in a previous year to produce a capsule. But
another species can easily encroach or simply intermix
enough to confuse the unwary. Be sure to track the
sporophyte down and locate its attached gametophyte. You
might find it belongs to a small pleurocarpous moss that is
weaving in and out among your acrocarpous cushion. This
sorting should be done with bryophytes that are moist
enough to be soft, but not soaked. Dry mosses are likely to
break before you can pull the gametophyte out from among
its trappings.
Wetting Agents
Assuming your specimens have not been collected in
the same day and have gotten dry and brittle, the first step
is to re-wet them before attempting to make a slide or even
examine them with the dissecting microscope. Dry
bryophytes are often brittle and will break easily if you
begin manipulating them without wetting them first.
However you wet them, we recommend watching them
with a dissecting microscope as the water moves through
the capillary spaces among the stems. It is a fascinating
display and is sure to grab the attention of first-time
viewers such as students.
Most bryophytes will wet up adequately by dipping
them in water or dropping water or misting on the desired
portion of the sample. Once the specimen has regained its
wet shape and is pliable, leaves can be removed by holding
the tips of the stems with a pair of forceps (can be ordinary
lab forceps if the specimen is not tiny) or a dissecting
needle (probe) and a second pair of microforceps should be
used to pull down on the desired leaf, being careful to hold
the leaf in a position close to the stem to get as much of its
base as possible. For smaller species, curved microforceps
often work best for holding the stems.
But some mosses simply don't wet well. In fact, some
bryophytes repel water and may even trap large air bubbles
that further keep them from getting wet. Members of the
Mniaceae are notorious for this, and Polytrichaceae can
be contrary as well if water cannot reach the leaf bases
easily. In particular, wetting agents help to avoid the air
bubbles trapped in leaf folds by reducing or eliminating the
surface tension of the water. Warm water can sometimes
actually increase the bubbles. Soap is a wetting agent, and
it doesn't take much. One drop in your dropper bottle is
likely to be more than needed. But beware, soap and the
other wetting agents, as well as heating, will usually kill the
bryophytes and destroy the cell contents.
One solution to getting some of these to get wet is to
drop them in hot (not boiling) water (Jewett 1913; Lucas
2009). I have to wonder if this distorts anything, and it
most likely melts waxes such as one might find on the
Polytrichaceae. But it does make most of them flexible
rather quickly, and lost wax is usually not a problem.
Some bryologists actually keep a hot plate nearby with hot
water while they work.
Koponen (1974) dips members of Mniaceae into 7090% ethanol, then into 2% KOH, ultimately washing away
the KOH with water. The specimens are ready for
examination in one minute and the chloroplasts are
destroyed, making other cell contents visible and the cell
walls a yellowish to brownish color. This is especially
helpful when the corners of the cells must be seen clearly.
A traditional wetting agent is one known by the
German word Pohlstoffe. This is a non-technical name for
a wetting agent (di-octyl sodium sulfosuccinate) available
from Fisher Scientific, known as Aerosol OT (Wagner
1981; Bryonet 23 July 2008); it is mixed in a 1:24:75 ratio
with methanol and water. Wagner suggested omitting the
methanol, finding that this modified mix brings leafy
bryophytes, dry capsules, and peristomes to turgidity
rapidly, virtually everything except thallose liverworts.
Schofield (1985) likewise suggested using only Aerosol OT
and water with a dilution of 1:100. It is named for Richard
Pohl (Diana Horton, Bryonet 19 September 1999) who
presented the formula as a softening agent for dried plant
parts (Pohl 1954).
Wagner recommends a half dropper of the 10%
solution in 50 ml of water in a dropping bottle. The
Aerosol OT can be difficult to obtain, especially if you are
not affiliated with an institution. A Google search only
located sites that sold it in huge quantities at costs of $500
or more. Wagner (Bryonet 11 May 2010) learned from his
students that the critical substance is also known as
docusate sodium, the active ingredient of stool softener!
Hence, it is available at the drugstore for about US $5.00
for 60 caplets (Figure 4). Wagner determined that one
caplet with a liquid center (100 mg docusate sodium), not
solid pills, in 25 ml of water works well as Pohlstoffe. The
carriers (glycerine, gelatin, propylene glycol, polyethylene
glycol) do not appear to leave any noticeable residue.
Soap (Tom Thekathyil, Bryonet 12 May 1210)
Another solution to wetting bryophytes is to use soap
or detergent as a wetting agent. Tom Thekathyil (Bryonet
12 May 1210) suggests diluted kitchen detergent. It doesn't
take much. One drop in your dropper bottle is likely to be
more than needed. A word of caution: Soap can destroy
the oil bodies of leafy liverworts! Warm water and
patience is a better approach.
2-2-5
Figure 4. Examples of stool softeners with docusate sodium.
Photos modified by Janice Glime.
Agral 600 (Tom Thekathyil, Bryonet 12 May
1210)
Tom Thekathyil also uses Agral 600 (horticultural
wetting agent). The latter kills the animal life that often
accompanies the bryophytes but does not seem to affect the
plants. This is useful to avoid introducing dermestids and
other hungry creatures into the herbarium.
Clearing Leaves (Rod Seppelt, Bryonet 13
May 2010)
I (Glime) have never tried clearing leaves – I wish I
had known about this for some of those dirty aquatic
species!
Lactic Acid
Lactic acid would have been helpful for those dirty
aquatic bryophytes. The lactic acid clears all the gunk from
the cells, making the walls much easier to see (Rod
Seppelt, Bryonet 13 May 2010). Rod Seppelt (Bryonet 13
May 2010) uses lactic acid to clear leaves. One drop on a
whole mount is sufficient (particularly if small), or with
leaves and sections. The lactic acid may also be added
under the cover glass of stems and leaves that have been
mounted moist, but not flooded. Gently warm the slide
using heat from an incandescent desk lamp. In the lab, if
you don't have an incandescent lamp, you can use a hot
plate, an alcohol burner, or even a candle, but you will need
to clean the carbon off the slide if you wave the slide
through the flame or place the slide above the flame. A
Bunsen burner is too hot and could result in boiling the
solution, a mishap to be avoided!
Unfortunately, lactic acid has its problems. It is
somewhat a health hazard if you make contact with it, but
less so than phenol, and it is not permanent on the slide.
Specimens need to be examined (and drawn if desired)
within a few days to weeks.
Water boils more quickly and suddenly than lactic
acid, so less water is better. One Bryonetter suggested that
2-2-6
a few air bubbles under the cover glass can be a useful
indicator of imminent danger. When the bubbles begin to
expand rapidly, it is time to remove the slide and let it cool
so it doesn't boil. If the solution reaches boiling, you will
most likely lose most of your dissected leaves and stems as
bubbles escape.
KOH or NaOH
Usually these methods will only require a few minutes
to clear the specimens. However, for especially dirty ones,
you may need to leave the specimen overnight to
clear. Potassium hydroxide (KOH) or sodium hydroxide
(NaOH) will also clear tissues – particularly if the material
is in contact with the air.
Stains can be used for a variety of purposes. They can
distinguish cell types, make pores visible, clarify cell walls,
make starch visible, and solve other problems in
distinguishing special structures. Most stains are readily
available, some are toxic, and others are household items.
Tom Thekathyil (Bryonet 13 May 2010) suggests
using household chemicals such as those provided by
Maier (2012). These include one drop of red or blue food
coloring in 30 mL of water, or for greater detail and
contrast, a mix of one or two drops each of red and blue
food coloring, five drops water, two to three drops white
vinegar, and three to five drops rubbing alcohol.
Chloral Hydrate
Chloral hydrate works well as a clearing agent, but
please read the discussion of its use in Chapter 2-4 of this
volume. It is a controlled substance and is dangerous to
your health.
If you should choose to use it, the following protocol,
developed for clearing parts of the flowering experimental
plant Arabidopsis thaliana, may be a useful start (Berleth
& Jurgens 1993). Substitute solutions for clearing can be
tried in place of the chloral hydrate – experiment:
1. Fix plant tissue in 9:1 parts ethanol:acetic acid. Use
vacuum infiltration to facilitate penetration of the fix
– approximately 2 hours at ambient temperature.
2. Wash tissue twice with 90% ETOH for 30 minutes
each wash.
3. Make solution of chloral hydrate or substitute in 30%
glycerol. (Note that another substitute might already
contain some glycerol.)
4. Add enough clearing agent (chloral hydrate or
substitute) to cover the tissue in an Eppendorf tube
(ca. 500 mL). Allow tissue to clear several hours.
5. Dissect tissue further if needed, using dissecting
microscope. Mount dissected, cleared plant parts in
chloral hydrate/glycerol or substitute under coverslip.
Seal slide with clear fingernail polish if desired.
Figure 5. Moerckia blyttii fresh plant. Photo by David
Wagner.
Dehydration
Usually specimens are air dried and this is adequate for
most species.
Some thallose liverworts require
preservation, but mosses rarely do. For higher quality
specimens, cleaned specimens can be dehydrated with a
series of ETOH (70, 90, 100%) (Mayfield et al. 1983).
Following the dehydration series, specimens are placed in a
1:1 ethyl alcohol:xylene solution, then transferred to 100%
xylene. Remove any remaining dislodged soil particles
with fine needles. The thalli can then be placed on glass
slides in a xylene-soluble mounting medium such as
Permount with coverslips that are weighted down with
small weights like nuts (of nuts and bolts) or metal washers.
Stains
For most observations, stains are not necessary. But
some things are simply too transparent or lack contrast.
The series of images of Moerckia blightii by David
Wagner (Figure 5-Figure 8) illustrate what stains can do to
aid visibility of the thallus structure.
Figure 6.
Moerckia blyttii cleared and stained with
methylene blue. Photo by David Wagner.
2-2-7
Etzold Stain (credited to Dr. Etzold) and W3A. The latter
is described (in German) at <http://www.mikroskopiebonn.de/_downloads/Arbeitsplan_W3Asim.pdf>.
Etzold Stain
Dissolve in 1L water:
Acetic acid (100 %): 20 ml
Fuchsin (bas.) 0.1 g
Chrysoidin
0.143 g
Astralblue
1.25 g
Color Results
non-ligneous cell walls: blue
ligneous cell walls, sclerenchym and xylem: red
Phloem: blue
Figure 7. Moerckia blyttii cleared and stained, grey scale.
Photo by David Wagner
Figure 8. Moerckia blyttii cleared and stained; gray scale
positive image converted to negative. Photo by David Wagner.
Staining Stems
Stems usually have specialized cells, including the
epidermis, the cortex, and often a central strand. Others
may have hydroids and leptoids and a second
distinguishable layer inside the epidermis. In some species,
natural colors distinguish the layers, but other
specializations may not be easily recognizable.
Triple Stains
Ralf Wagner (pers. comm. 2012) suggests two triple
stains that can be used to distinguish cell differences, the
Kawai Stem Staining Techniques
Kawai did extensive studies on stem sections using a
variety of dyes (Kawai 1971a, b, c, 1974, 1975, 1976,
1977a, b, 1978, 1979, 1980a, b, 1981, 1982, 1989, 1991a,
b; Kawai & Ochi 1987; Kawai et al. 1985, 1986) (Figure 9Figure 48). He cut stems in 5 or 10 µm, even 15 µm
sections (Isawo Kawai, pers. comm. 5 October 1989).
Most of the information we have is the result of personal
communication and a set of images he sent to me (Glime)
many years ago. The effectiveness and time required
varied among species and even within a species, perhaps
indicating differences in age of the tissue or habitat where
it grew.
For his early studies on mosses (Hypnaceae,
Thuidiaceae), Kawai (1971c, 1975, 1976) rehydrated the
mosses by boiling them for half an hour to an hour in water.
He then used a standard technique of ethylalcoholbutylalcohol-parafin for fixation. Sections were usually 5
µm thick.
As his work progressed, he experimented with various
methods of staining. In early studies, he used acid fuchsin,
fuchsin, fast green, and methyl green to stain members of
Bartramiaceae, Dicranaceae, Entodontaceae, and
Fissidentaceae (Kawai 1971).
In some cases
(Amblystegiaceae
sensu
lato,
Dicranaceae,
Fontinalaceae,
Hedwigiaceae,
Lembophyllaceae,
Leucodontaceae,
Meteoriaceae,
Neckeraceae,
Pterobryaceae, Trachypodiaceae), he used just gentian
violet and acid fuchsin (Kawai 1977b, 1978, 1979). In
others (Amblystegiaceae, Bartramiaceae, Dicranaceae,
Hypnaceae, Leucobryaceae) he stained with gentian
violet, acid fuchsin, and potassium iodide, using 5 µm
sections (Kawai 1980a, b, 1981, 1982). As part of his
experimentation with methods, he used 15 µm sections
with the Bryaceae (Kawai & Ochi 1987).
Some mosses were much more resistant to the stains.
In particular, members of the Polytrichaceae and
Fontinalaceae were difficult to stain so that cell types
could be seen clearly (Kawai, pers. comm. 5 October 1989).
Kawai et al. (1985, 1986) ultimately developed a lengthy
and more complex protocol that gave satisfactory results.
Even this differed between species within the family.
For Polytrichum commune, Kawai et al. (1985) tried
three methods. 1) Aniline Blue-Eosin-Methyl Green
Method: They placed the moss in a solution of aniline
blue and eosin for 48 hours, followed by washing and a
second solution of just eosin for another 48 hours. Finally,
2-2-8
the preparation was washed again and placed in a solution
of methyl green for another 48 hours. After washing the
stems were cut in 15 µm sections with a cryo-microtome
and mounted in gum arabic. 2) Janus Green-EosinMethyl Green Method: The specimen was soaked in a
solution of Janus green and eosin for 48 hours, washed,
and soaked another 48 hours in just eosin. The specimen
was washed again and soaked in methyl green for 48
hours before the final washing, sectioning, and mounting.
3) Gentian Violet+Congo Red-Eosin-Methyl Green
Method: The specimen started in a solution of gentian
violet and Congo red for 32 hours. As in the other
procedures, it was washed and soaked in eosin, this time
for 40 hours. Finally it was washed and placed in a
solution of methyl green for 32 hours, washed, sectioned,
and mounted.
For Pogonatum contortum, Method 1 was successful,
but specimens were soaked in each solution for 32 hours,
except for 40 hours for just eosin (Kawai et al. 1985). For
Rhizogonium and Mnium, Method 1 was successful, but
specimens were soaked in each solution for 72 hours. For
Fissidens, Method 2 was successful, but specimens were
soaked in each solution for 36 hours.
In general, Kawai used the following concentrations:
eosin 0.2 g per 100 cc
methyl green 0.005 g per 100 cc
Figure 9-Figure 48 illustrate the responses of a variety of
species in various soaking times.
Figure 11. Fontinalis gracilis stem cross section stained
with aniline blue for 5 minutes. Photo by Isawo Kawai.
Figure 12. Bryoxiphium sp. stem cross section stained with
aniline blue for 1 hour. Photo by Isawo Kawai.
Figure 9. Fontinalis antipyretica stem cross section stained
with 0.005 g per 100 cc methyl green for 10 seconds, then
stained with 0.2 g per 100 cc eosin for 50 minutes. The bluegreen/green color clearly shows the inner layer of "epidermal"
portion of the stem. Photo by Isawo Kawai.
in aniline blue for 20 minutes. Photo by Isawo Kawai.
Figure 10. Polytrichum sp. stem cross section. The cortex
cell walls are blue-green from methyl green. The hydrome cells
are violet-brown. Photo by Isawo Kawai.
with aniline blue for 1 hour. Photo by Isawo Kawai.
2-2-9
Figure 19. Hylocomium sp. stem cross section stained with
aniline blue for 3 hours. Photo by Isawo Kawai.
Figure 20. Hypnum sp. stem cross section stained with
with aniline blue for 1 hour. Photo by Isawo Kawai.
Figure 21. Polytrichum sp. stem cross section stained with
aniline blue for 2 hours. Note the cell inclusions in these cortex
cells. Photo by Isawo Kawai.
Figure 18. Fontinalis hypnoides stem cross section stained
aniline blue for 2 hours. Photo by Isawo Kawai.
2-2-10
Figure 23. Rhizogonium sp. stem cross section stained with
with aniline blue + eosin for 3 hours. Photo by Isawo Kawai.
in aniline blue + eosin for 1 hour. Photo by Isawo Kawai.
in aniline blue + eosin for 1 hour. Compare this to the previous
picture (Figure 24) to see differences that can occur under the
same staining protocol. These differences may relate to age of the
tissues or possibly the habitat. Photo by Isawo Kawai.
Figure 28. Hylocomium sp. stem cross section (5 µm thick)
stained with aniline blue + eosin for 2 hours. Photo by Isawo
Kawai.
2-2-11
Figure 32. Thuidium sp. stem longitudinal section stained
aniline blue + eosin for 3 days. Photo by Isawo Kawai.
Figure 33. Bryoxiphium sp. stem longitudinal section
stained with eosin for 2 hours and methyl green for 30 seconds.
Photo by Isawo Kawai.
Figure 30. Thuidium sp. stem cross section stained with
aniline blue + eosin for 2 hours. Photo by Isawo Kawai.
aniline blue + eosin for 3 days, washed, stained with eosin 3
more days, then stained with methyl green. Photo by Isawo
Kawai.
Figure 34. Bryoxiphium sp. stem cross section stained with
eosin for 2 hours and methyl green for 30 seconds. Photo by
Isawo Kawai.
2-2-12
Figure 35. Fontinalis gracilis stem longitudinal section
stained with eosin for 1 hour, washed, then stained with methyl
green for 30 seconds. Photo by Isawo Kawai.
Figure 38. Hypnum sp. stem longitudinal section stained
with eosin for 1 hour, then stained with methyl green for 30
seconds. Photo by Isawo Kawai.
Figure 36. Hylocomium sp. stem cross section stained with
eosin for 1 hour, then with methyl green 1 minute. Photo by
Isawo Kawai.
stained with 0.005 g per 100 cc methyl green for 10 seconds,
then stained with methyl green + 0.2 g per 100 cc eosin for 15
minutes. Photo by Isawo Kawai.
Figure 37. Hypnum sp. stem cross section stained with
eosin for 1 hour, washed, then stained with methyl green for
0.5-1 minutes. Photo by Isawo Kawai.
stained with methyl green + 0.2 g per 100 cc eosin for 15
minutes. Photo by Isawo Kawai.
stained with methyl green + 0.2 g per 100 cc eosin for 1 hour.
stained with 0.005 g per 100 cc methyl green for 10 seconds,
then stained with methyl green + 0.2 g per 100 cc eosin for 1
hour. Photo by Isawo Kawai.
Figure 43. Polytrichum sp. 10 µm stem cross section stained
with 0.01g per 100 cc methyl green for 50 seconds, then 0.3 g per
100 cc eosin was added for 2 hours, then washed with water.
2-2-13
0.01g per 100 cc methyl green for 50 seconds, then stained with
0.3 g per 100 cc eosin for 2 hours. Photo by Isawo Kawai.
0.01g per 100 cc methyl green for 3 minutes, then stained with
eosin for 2 hours. Photo by Isawo Kawai.
eosin for 1 hour, then stained with methyl green for 2 minutes.
2-2-14
eosin for 2 hours, washed, then stained with methyl green for 1
minute. Photo by Isawo Kawai.
Figure 48. Thuidium sp. stem cross section stained with
eosin for 2 hours, washed, then stained with methyl green for 1
minute. Photo by Isawo Kawai.
Acid Fuchsin
Acid fuchsin has been used to stain a variety of plant,
animal, and fungal tissues. Kawai (1980b) used acid
fuchsin, along with I2KI and gentian violet to distinguish
the internal anatomy of stems in the Leucobryaceae.
Using 5 µm sections, he was also able to examine the
structure in Atrichum undulatum, Bartramia pomiformis,
Dicranum nipponense, Leucobryum neilgherrense, and
Hypnum plumaeforme (Kawai 1981).
Aniline Blue (Kawai & Glime 1988)
Kawai used aniline blue to stain several species,
including Fontinalis spp. (pers. comm. 5 July 1988),
Polytrichum commune, and Pogonatum contortum
(Kawai et al. 1985). It stained the epidermal (outermost
layers of stem) red and those just inside the red ones were
stained blue.
1. Place moss in solution of aniline blue and eosin for
48 hours. The hydrome cell walls stain violet-brown.
2. After washing, place the moss in eosin for absorption
for 48 hours to stain epidermal cell walls and leptome
red.
3. Wash again and place moss in solution of methyl
green for another 48 hours to stain cell walls of
cortex blue-green.
Congo Red (Kawai & Glime 1988)
1. Place leafy gametophyte into solution of gentian
violet and Congo red for 48 hours to stain hydrome
cell walls violet-brown.
2. Wash moss and place in solution of eosin for another
48 hours to stain cell walls of epidermis, cytoplasm of
leptome, and chloroplasts red.
green for another 48 hours to stain cortex cell walls
blue-green.
Eosin
Eosin is a red dye that stains cytoplasm. It is watersoluble and thus can be used to follow water movement
through plants. It has been used in the tracheophyte
Arabidopsis sp. to indicate photodamage to the
photosynthetic apparatus (Havaux et al. 2000).
Kawai (pers. comm. 8 July 1989) used eosin as one of
the stains to distinguish cells in Fontinalis antipyretica.
This stains the outer cells of the stem ("epidermis") (Figure
9) and the cell walls of the cortex red. Eosin likewise
stained the cytoplasm of the leptom and the chloroplasts
red. As noted earlier, this stain works well in most
bryophytes to stain cell walls and cytoplasm red.
Fast Green
Fast green is the green dye used in food coloring, but
it is known to have tumorogenic effects. It is a protein
stain and is one of the stains used by Kawai (1971).
Fuchsin
The dye fuchsin is a biological stain that is produced
by oxidation of a mixture of aniline and toluidine,
producing a brilliant bluish red. Kawai (1971) used it to
stain bryophyte stems.
Gentian Violet (=Crystal Violet)
The color of stain by gentian violet depends on the
acidity. At pH 1.0, the dye is green, but in an alkaline
solution it is colorless.
Kawai (1980b) used gentian violet, along with acid
fuchsin and I2KI to distinguish structures within the stems
in members of the Leucobryaceae.
Janus Green
Janus green is a vital stain that changes color based
on the level of oxygen in a cell (Wikipedia 2012). Kawai
(pers. comm.) has used it in combination with other stains
to stain the hydrom of moss stems.
Methyl Green
Isawo Kawai (pers comm. 8 July 1989) used 0.005 g
per 100 cc of methyl green for 10 seconds to stain cells in
10 µm sections of the stem of Fontinalis antipyretica
(Figure 9). This was followed by eosin (0.2 g per 100 cc)
added to it. This mix was allowed to stand for 50-60
minutes, then washed for observation. Eosin stained the
outer cells of the stem red and methyl green stained those
just inside the outermost layers a blue-green color (Figure
9). The central tissue did not stain with this combination.
1. Place leafy moss in solution of Janus green and eosin
for 48 hours to stain hydrome cell walls violet-brown.
2. After washing, place moss in solution of eosin for
further 48 hours to stain cell walls of epidermis,
cytoplasm of leptome, and chloroplast red.
green for another 48 hours to stain the cortex cell
walls blue-green.
Kawai and coworkers (Kawai et al. 1985; Kawai, pers.
comm. 5 October 1989) found that the leaf cell walls of
Polytrichum sp. (Figure 10), Fissidens sp., and Bartramia
sp. stained blue-green with methyl green, but the cell walls
of several species of Fontinalis leaves (Kawai, pers. comm.
5 October 1989) would not stain with methyl green.
Leaves
I2KI – Lugol's Solution (Kruijer & Klazenga
1994)
Kruijer and Klazenga (1994) consider methylene blue,
a common Sphagnum stain, to be somewhat problematic
for other leaves, sometimes staining too darkly. Instead,
they recommend staining with a diluted solution of iodinepotassium iodide (I2KI), or Lugol's solution (Johansen
1940). This is the well known stain for starch, causing it to
turn blue to purple to nearly black. But it can also stain
cellulose if tissues are first hydrolyzed with sulfuric acid
and hemicellulose if hydrolyzed with hydrochloric acid.
Kruijer and Klazenga used I2KI successfully on leaves and
cross sections of members of the Hypopterygiaceae and
the genus Dicranoloma. Cell walls became brighter, but
remained nearly colorless except for the middle lamella,
which sometimes became bright yellow.
KOH (Zander 1989, 1993)
KOH in concentrations from 2% to saturated will stain
cell walls of many mosses. It can be used on whole leaf
mounts and on sections. In the Pottiaceae the resulting
colors can be used as diagnostic characters. Zander (1993)
uses it to rehydrate mosses as well. The KOH should not
be kept in glass dropper bottles because it reacts with the
glass to form a precipitate. If the specimen will later be
mounted with an acidic mountant, add a drop or two of
dilute HCl to the specimen.
KOH is useful in distinguishing between genera in the
Pottiaceae (Zander 1993). For example, the lamina color
reaction in Tortula and Ganguleea is yellow, whereas in
Syntrichia, Dolotortula, Chenia, Hilpertia, Sagenotortula,
Stonea, and Hennediella it is red, and in Saitoa, deep red
(Zander 1989).
Safranin O / Fast Green (Rod Seppelt, Bryonet
15 August 2012)
Rod Seppelt (Bryonet 15 August 2012) considers this a
good general stain for plant sections. It works well for
bryophytes on specimens that have been fixed and
embedded and on sectioned material. Bill and Nancy
Malcolm (2006) have used this combination to obtain highcontrast color effects. The technique is somewhat timeconsuming, requiring a schedule of dehydration and
rehydration. They suggest a quicker option using toluidine
blue. If it is used simply to clear the cells, then the
hydrolyzation step is unnecessary.
Lisa Op den Kamp (Bryonet 4 October 2012) also
uses safranin. She applies this directly to the leaves or
capitula of Sphagnum, then washes them in water, all
2-2-15
before cutting the Sphagnum to make the desired sections.
Safranin normally dyes lignin red; although Sphagnum
doesn't have typical lignin, safranin stains the lignin-like
compounds in the tissues. She has kept the solution for 12
years and it still works well.
Sphagnum Stains
In particular, Sphagnum leaves typically need to be
stained for the pores to be visible. Rudi Zielman (Bryonet
4 October 2012) considers there to be four Sphagnum
stains:
aniline blue
methylene blue
gentian violet (=crystal violet)
toluidine blue O
These can be applied in two ways: supply a bit of it
directly in a few drops of alcohol or water or make a stock
solution based on alcohol or water.
To enhance the pores on Sphagnum leaves, Rod
Seppelt (Bryonet 13 May 2010) suggests toluidine blue,
aniline blue, or methylene blue. A drop or two in 100 ml
of water should be sufficient.
Schofield (1985)
recommends methylene blue, gentian violet, or crystal
violet in a 1-2% aqueous solution. If the stain is very dark,
simply dip the moss in quickly and then rinse it in clear
water. If it gets too much stain, you will see even less than
with no stain. Be careful – these stains also stain fingers
and clothing! If you don't have the standard stains, try
experimenting. We wonder if beet juice would work. It
might need a bit of vinegar to make it colorfast for
permanent mounts.
Methylene Blue (Kruijer & Klazenga 1994;
Wagner, Bryonet 11 May 2010)
When staining Sphagnum pores, it is important not to
stain too heavily. Kruijer and Klazenga (1994) use a 1-2%
aqueous solution of methylene blue. Or, place a drop of
full strength dye on a slide or in a Syracuse watch glass.
Dip the Sphagnum branch quickly into the dye to cover the
branch, then dip the branch into clean water to wash the
dye off. Don't allow the branch to remain in the dye. After
washing, the moss should be ready for viewing.
David Wagner (Bryonet 11 May 2010) brings us a
simple solution for staining Sphagnum, a contribution
from one of his students. Since methylene blue is used as
an antibiotic for aquarium fish for hatching eggs or getting
rid of fungal infections, it is readily available at tropical
fish stores. A half ounce bottle (ca 12 ml) of VERY
concentrated methylene blue is only about US $4.25 and
will be a lifetime supply.
Crystal Violet/Gentian Violet
Crystal violet, also known as gentian violet or methyl
violet 10B, is the compound hexamethyl pararosaniline
chloride, or pyoctanin(e), and is a triarylmethane dye.
Adam Hölzer (Bryonet 4 October 2012) reports that he
can see even the pores of Sphagnum obtusum (Figure 49)
very well with crystal violet. He dissolves some powder in
about 50 ml of distilled water with the addition of some
alcohol to preserve it. He adds new alcohol occasionally to
compensate for evaporation. He puts the moss leaves in a
drop of water. Then uses his forceps to dip into the solution
and transfer only a small drop into the drop of water. He
covers the drop with a cover glass. The color stains the
2-2-16
cellulose of the leaves Excess stain can be removed by
adding water to one side and drawing off the solution on
the other with tissue paper. The 50 ml of solution will last
for several years even if you use it every day. Stains on the
desk can be cleaned with alcohol.
cheaply (e.g. items 261098492176 and 261107216623).
But he cautions that for non-aqueous permanent mounts,
these stains are not suitable. Instead, Bismark Brown
provides a nice stain.
Reproductive Structures
Iron Haematoxylon / Fast Green (Rod Seppelt,
Bryonet 15 August 2012)
This stain works very well to show archegonia and
spermatogenous cells in antheridia (Rod Seppelt, Bryonet
15 August 2012).
Bulbils and Spores
Figure 49. Sphagnum obtusum stained for pores. Photo by
Ralf Wagner.
Crystal violet and gentian violet solutions can be used
to fill well-rinsed and dried felt-tip pens (Joannes (Jan) A.
Janssens, Bryonet 4 October 2012). These pens can be
used in the field to stain Sphagnum that has been squeezed
somewhat dry.
Toluidine Blue O (Rod Seppelt, Bryonet 15
August 2012)
Rod Seppelt (Bryonet 15 August 2012) considers this
to be the most useful stain for general tissue differentiation
in fresh material, but it is not useful for permanent mounts.
It can help to distinguish the ventral row of leaves in
liverworts. It also will reveal the pores and stem leaves in
Sphagnum.
Simple method:
0.2%-0.25% toluidine blue O in water (be sure it is O),
or 1 drop in 10 drops of water
Stain moss in solution for 10-30 seconds, place on
slide, apply cover glass, and examine (without washing
excess stain away). If too dark, dilute the stain further
before use, or wash the moss quickly to remove some of
the excess.
In vascular plants, its multiple color responses can
indicate tissue type:
phloem green, xylem blue,
parenchyma purple, lignified tissue of bundle caps pale
whitish-green. Similar color distinctions may occur in
bryophytes. Unfortunately, the color is not permanent.
More complex recipe:
0.610 g KH2PO4
0.970 g K2HPO4
0.050 g Toluidine Blue O
In 100 ml distilled water
Des Callaghan (Bryonet 4 October 2012) likewise
recommends Toluidine Blue O and Safranin O for
Sphagnum. Simply dip the branch in the stain and it works
almost immediately. You can find the stains on eBay
Fluorescence and Fluorescent Dyes
(Nordhorn-Richter 1988)
Gisela Nordhorn-Richter (pers. comm.) discovered the
fluorescence of bulbils in Pohlia when a microscope
salesman visited her institution. No one was visiting the
display and she felt sorry for the salesman, so she took
some of her specimens to look at them. She was amazed at
the ease of finding bulbils with the fluorescence technique.
Preparation of bryophytes for fluorescence microscopy
is mostly what not to do. They can be prepared on a slide
with water or as permanent slides (Nordhorn-Richter 1988).
However, some of the embedding materials have phenolic
compounds as preservatives or may have a synthetic resin.
These produce fluorescence that interferes with seeing the
bryophyte structures. Air bubbles are another potential
problem because they can scatter the light. Dry plants can
only be rewet once because the membranes typically are
destroyed by drying. When the plants are rewet, water
soluble substances leak from the cell. When they dry once
again, the water-soluble fluorescing substances disappear,
ending fluorescence.
In the dried condition, fluorescing substances of
bryophytes are very stable, with rhizoid bulbils of Pohlia
that are more than 100 years old still exhibiting brilliant
fluorescence. Chlorophyll, on the other hand, loses its
fluorescent ability upon drying.
To hide the fluorescence of chlorophyll, which can
interfere with fluorescence of other substances, a
suppression filter of 650 nm can absorb its red fluorescence
(Nordhorn-Richter 1984a, b, 1985a, b, 1988).
Alternatively, the chlorophyll can be extracted by 80%
acetone or DMSO without interfering with other
fluorescent substances.
The fluorescence technique for bryophytes permits one
to find rhizoid gemmae hiding in a sandy substrate
(Nordhorn-Richter 1988).
Live spores exhibit red
fluorescence, permitting estimation of vitality that can be
quantified with a fluorescence spectrophotometer (Figure
50; Ridgway & Larson 1966; Paolillo & Kass 1973; Genkel
& Shelamova 1981). Phenolic acids, including Sphagnum
acid (Tutschek 1975), lignin-like compounds in cell walls
(Lal & Chauhan 1982; Nordhorn-Richter 1984a, 1985),
peristome structure (Nordhorn-Richter 1985b), and papillae
(Nordhorn-Richter 1984b) become visible. Even small
bryophytes can be found by using a UV light (366 nm) at
night! (Nordhorn-Richter 1983). Gambardella et al. (1993)
used fluorescence microscopy to examine the cytoskeleton
of the columella in Timmiella barbuloides.
Animal
tissues exhibit only secondary fluorescence, making it
possible to distinguish between animal galls and bryophyte
propagules (Nordhorn-Richter 1988).
2-2-17
Staining Liverwort Capsules (Von Konrat et al.
1999)
Von Konrat et al. (1999) devised a technique to
examine the multiple layers of the capsule wall of
liverworts. First the layers need to be separated using a
pectinase preparation. Then the layers need to be cleared
and stained to make the details easier to see.
They recommended doing all the treatments on the
same slide – results were less satisfactory when the
specimen was moved from one reagent to another.
Solutions can be removed between treatments by using
filter paper cut into strips. The capsule was mounted on a
coverslip and the fully stained capsule was mounted
between two coverslips so that both surfaces could be
examined.
1. Treat with FAA for 24 hours or until decolored
Figure 50. Spores of Fontinalis squamosa showing spores
in white light on left and fluorescing under UV light on right.
Note that the living spores show up as red under fluorescence,
whereas dying and dead spores are yellow or invisible. Photos by
Janice Glime.
Shellhorn et al. (1964) demonstrated that both fresh
and fossil pollen could be detected with fluorochromes,
with better results if acridine orange was added to enhance
detail.
Ridgway and Larson (1966) extended the
fluorescence technique to provide better viewing of the
features of the hornwort Anthoceros. The images of spores
of Fontinalis squamosa demonstrate that the use of
fluorescence microscopy can help to distinguish living
from dead spores in mosses (Figure 50). The yellow
fluorescence in the image suggests that the exine is
fluorescing, as it did in pollen (Ridgway & Larson 1966).
Stains can provide one with the ability to see structures
using fluorescence microscopy. Brandes (1967) explained
the use of acridine orange as a vital stain for use with
fluorescence microscopy of protonemal pro-buds and buds.
The stain moves to the cytoplasm, combining with the
RNA. This technique shows the increase of cytoplasmic
RNA immediately after the induction of the pro-buds.
Hence, kinetin-induced buds, as well as non-induced
branches, can be detected ten hours after the beginning of a
kinetin treatment.
Fluorescent dyes can have various purposes, including
using them as growth markers in the field (Russell 1988).
Fluorochrome
3,3'Dihexyloxacarbocyanine
iodine
[DiOC6(3)] can be used to locate selectively the fungal
hyphae among the rhizoids of bryophytes (Duckett & Read
1991).
Ascomycetous hyphae are visible when
concentrations of 0.01-5 µg ml-1 are used, whereas to see
Basidiomycetes that form endophytic associations, the
concentration needs to be at least 50 µg ml-1. Some fungi,
such as VA fungi in liverworts, do not stain with
fluorochrome at any concentration. Others require a much
lower concentration than these. One advantage to this
method is to recognize the extent of the fungal hyphae in
the association.
FAA (Formalin-Acetic-Alcohol)
(100 ml)
Ethyl alcohol
50 ml
Glacial acetic acid 5 ml
Formaldehyde (37-40%)
10 ml
35 ml
Distilled H2O
2. Rinse in water three times.
3. Clear partially with 80% lactic acid at 60°C for 30-60
minutes in container saturated with water vapor.
4. Wash again at least three times in water.
5. Add enough 1% (v/v) pectinase preparation of
Aspergillus niger in water to cover specimen. Let
stand for a maximum of 1 hour at 37°C with slide in
container saturated with water vapor.
Catalog # P-9179
Sigma Chemical Co.
St. Louis, MO, USA
6. At this stage, you should be able to find the cell layers
separated or at least tissue fragments from internal
layer separated from the epidermal layer, permitting
adequate comparisons.
Longer digestion causes
digestion of the tissue and thus digestion should stop
after 1 hour even if tissues are not separated.
7. Rinse with water three times.
8. Add 1 drop of water and 3.5% sodium hypochlorite
(household bleach) for 30-120 sec or until capsule
becomes nearly transparent.
(Monitor under
dissecting microscope.)
9. Rinse with water three times for 30-60 sec each time.
10. Add dye for 60-120 seconds, depending on dye (see
Table 1 below).
11. Rinse again for 60 sec in water.
12. Examine capsules in water or glycerol. Water can
cause surface tension problems and material may
scatter, making glycerol preferable (Zander 1997).
13. If necessary, gentle tapping or squashing with a pair
of fine forceps may help to separate the internal layer.
14. Mountants may include Aqueous Mountant or
glycerol in glycerin jelly (Zander 1997). Hoyer's
solution is not suitable because the dye will fade.
2-2-18
Table 1. Von Konrat et al. (1999) tested coloration methods on the cell walls of the leafy liverwort Frullania.
Internal Layer
Stain
Epidermal Layer
Cell walls
Thickening
Cell walls
Thickening
Alcian blue
(0.02% w/v in water)
blue
+
blue
++
blue
+
blue
++
Autofluorescence
blue
+
–
blue
+
–
Bismark brown Y
(1.0% w/v in 5% w/v
aqueous phenol soln)
orange/brown
orange/brown
orange/brown
orange/brown
+
+
+
+
Calcofluor white
blue
+
–
blue
+
–
Methylene blue
blue
+
blue
+
blue
+
blue
+
p–Nitrobenzenediazonium
tetrafluoroborate
(0.5% w/v in 0.1 M sodium
phosphate buffer
pH 7.0 for 10 min at 4°C)
Ruthenium red
(0.02% w/v in 1% w/v aqueous
soln ammonium acetate)
Toluidine blue O
(0.05% w/v on sodium
benzoate buffer pH 4.4)
orange
orange
–
+
–
+
–
red
++
–
red
++
pink–purple
pink–purple
–
+
–
+
Nile blue A
(0.01% in water)
–
–
–
–
Phloroglucinol-HCl
(1 ml 2% w/v in 95% v/v
aqueous ethanol +
2 ml 10M HCl
–
–
–
–
Sudan red 7B
–
(0.1% w/v in 50% v/v
polyethylene glycol +
45% v/v glycerol + 5% v/v water)
–
–
–
pH Testing (Zander 1980; Long 1982)
Lichenologists are quite familiar with testing pH
reactions, but this technique has not been widely used on
bryophytes. Zander (1980) used pH responses (acid-base
color reactions) to separate Triquetrella californica from
Didymodon fallax var. reflexus and to remove
Bryoerythrophyllum calcareum and B. inaequalifolium
from the genus Barbula. Long (1982) similarly tested four
species of Pottiaceae and was able to distinguish them on
the basis of color change. He used concentrated HCl, 10%
KOH, concentrated nitric acid, and 2:1 concentrated
H2SO4, obtaining, respectively, the following results:
Bryoerythrophyllum wallichii – pale brown, red-brown,
red-brown, dark red-brown
Bryoerythrophyllum caledonicum – pale greenish-brown,
red-brown, red-brown, dark red-brown
Leptodontium flexifolium – green, orange, red, brown &
green
Paraleptodontium recurvifolium – green, orange, red, dark
brownish-green
Weak Alkali (Lane 1978)
Lane (1978) used a saturated solution of sodium
bicarbonate (Hill 1976) in distilled water (weak alkali, final
~pH 10) to effect color change in red-pigmented
Sphagnum. The branches or capitula were flooded by
pipette, then permitted to stand for 1-2 minutes (Lane
1978). He then permitted the flooded branches to dry
overnight, compared them to known specimens again, and
flooded them with a weak acid (e.g. vinegar) of ~pH 3 to
check for color change reversibility. Of the 17 species
tested, Lane found that there was no color change in
subgenera Rigida, Subsecunda, or Cuspidata, although
Subsecunda became redder. Sphagnum magellanicum
(subgenus Sphagnum) became dark brown-black.
Sphagnum wulfianum (subgenus Polyclada) became
chocolate brown. The nine species in subgenus Acutifolia
all turned blue or dark blue. The test works equally well on
fresh, freshly dried, and herbarium material.
Cleaning Up Stains
Spilled stains are hard to remove. David Wagner’s
experience testing kitchen cleaning agents for removing
stains from floors or bench tops has found “Bar Keepers
Friend”™ with oxalic acid is better than most.
Leaf Removal and Making Slides
For identification, cells, margins, costa, and insertion
of leaves must be seen clearly. In some cases, especially
leafy liverworts, these can be seen by making a slide of the
branch or stem intact. But for most mosses, it is too
difficult to see everything that is needed. Removing a leaf
from a moss is usually a necessity to attain this clarity.
There are a number of publications on preparing slides for
viewing bryophytes (Murray 1926). I have extracted from
these what works for me:
1. First moisten the moss by placing it in a beaker of
water.
2. Place a stem on a glass slide and strip the leaves by
pulling them downward from the tip with a pair of
microforceps while holding the tip of the branch or
stem with another pair of forceps. Alternatively, you
can run the convex side of a pair of curved
microforceps down the stem to break off leaves.
Some bryologists remove leaves by running a
dissecting needle down the stem while holding the tip
with forceps on a glass slide. Still others (Lucas
2009) use a spear point to run down the stem to
remove leaves. Lucas points out that the spear tip
tends to leave other structures such as paraphyllia on
the stem where they are more easily observed.
3. Remove most of the branches from the portion of
the stem you will observe on the slide (Lucas 2009)
so that the coverslip can flatten the stem better for
easier viewing. But you will also need to compare
branch and stem leaves, which differ in some species.
4. Put a drop of water on the leaves and/or stems and
spread them out so some are dorsal and others ventral
in position.
5. Hold the coverslip by its edges and lower one side of
the coverslip gently with a needle or forceps to avoid
trapping air bubbles (Figure 51). If you drop the
coverslip straight down, there will be no chance for
bubbles to escape. If the stem is bulky and the leaves
small, you might want to put them on separate slides.
Figure 51. Technique for making a slide with minimal air
bubbles. Drawing by Janice Glime.
2-2-19
6. If the coverslip is floating, remove some of the water
by touching a paper towel edge to one coverslip edge.
If there is not enough water, add water to the edge of
the coverslip with a dropper. Too much water will
allow your images to move about and wiggle, making
examination difficult. Too little will cause the water
to draw around the specimen and cause light
distortions.
7. Examine with the compound microscope. The
magnification depends on the size of the specimen
and what you are trying to see. It is usually best to
locate the specimen and focus on 40X or 100X, then
move to 400X when more detail is needed.
8. To see papillae, decurrencies, projecting costa tips,
and perhaps other surface features, you need to see
the leaf in side view, so it is best to observe the leaves
that remain on the stem for these features. Most other
features are best seen on detached leaves that are
more or less flattened by the coverslip. Look around
and observe several of the leaves.
Sectioning
It seems that bryologists have developed a number of
methods for sectioning bryophytes (e.g. Singh 1942; Foster
1977; Nishimura 1997). Nevertheless, Sean Edwards
(Bryonet 30 July 2002) points out that bryologists have
tended to avoid cutting sections of moss leaves for several
reasons:
1. Microtome sectioning involves some considerable
delay owing to the various preparations required
(moreover, microtomes are often not available,
especially to amateurs, when needed).
2. Pith sectioning is unsatisfactory because of the
difficulty in controlling section thickness, and in
separating the pith debris without damaging the
sections.
3. In both microtome and pith sectioning it is almost
impossible to be certain of the exact part of the leaf
from which the sections were taken.
4. In both methods of sectioning, considerable care and
time are needed to maintain a suitable cutting edge.
Nevertheless, there are several methods used by
bryologists for making sections of stems and leaves (e.g.
Singh 1942; Frolich 1984; Nishimura 1997). One is to
place the stem with leaves on a dry glass slide and chop,
like cutting parsley! The idea is that with lots of cuts, some
of them will yield a usable section.
Razor Blades
Razor blades are the standard tool for cutting sections.
Hutchinson (1954) recommends use of a normal razor
blade that is divided into four sections. The blade should
be placed between pieces of blotting paper and broken
down the center the long way. Each of these pieces is
broken again perpendicular to the previous break. She
found she could use used blades because only the sharp
points are needed. The blades can even be broken again
when the points become dull.
Cutting Techniques
In the many techniques that create sections, placement
of the sections is important. Once the specimen sections
are in a drop of water on the slide, Hutchinson (1954)
2-2-20
recommends stirring the water to distribute the specimens,
while looking through the eyepiece of a dissecting
microscope. Be sure the water is not sufficient to exceed
the area of the coverslip when it is applied or you will use
the smallest, hence the best, specimens. As the slide begins
to dry, add 5% glycerine at the edge of the coverslip. If the
best specimens need to be moved to another slide, you can
use a dental applicator (Figure 52) dipped in a 5% solution
of glycerine. When placed over the desired section, this
combination will lift it up. The applicator can be dipped
into a drop of the same solution on the new slide and the
section shaken off.
7. If leaves curl, soak in solution A at room temperature
until solution reaches consistency of glycerin.
8. Remove leaves and touch to filter paper to remove
excess liquid.
9. Transfer blotted moss to molten B and proceed from
#1.
10. Transfer cut sections with adhering wax to water with
small amount of wetting agent if need to keep from
floating. Taylor prefers enough water to cover bottom
of Syracuse watch glass.
11. Sections can be transferred by tapping slide on rim of
watch glass.
12. Polyethylene glycol is not compatible with gelatin, so
sections should stay in water until wax completely
dissolves – a few minutes in warm water.
13. Remove sections and put in dilute glycerin onto slide.
Cutting Block (Flowers 1956)
Flowers (1956) used a 2x2x15 cm cutting block made
of soft wood. She then made a jellyroll arrangement of the
bryophyte in tracing paper (a thin paper):
Figure 52. Dental Disposable Micro-Applicators. White is
superfine, yellow is fine. Photo modified from AliExpress.
Wax Mounts (Taylor 1957)
Taylor (1957) found a different solution to positioning
leaves and stems for cutting. He first coats them with
water-soluble wax. These include Carbowax and some
kinds of crayons.
Taylor makes two solutions: Solution A is 20%
aqueous polyethylene glycol 600 with a small quantity of
Quarternary amine disinfectant to prevent development of
fungi in permanent mounts. Solution B consists of
polyethylene glycols 1540 and 4000, which can be used
alone or in combination. However, 1540 alone may be too
soft, and 4000 too crumbly.
1. Place solution B on a slide and melt.
2. Place a piece of stem in molten drop to cover stem.
The drop needs to be thick enough to support the
blade during cutting.
3. Cool wax for ~1 minute with slide on cool metal
surface.
4. Use quarter of razor blade to trim drop at one end to
point where sectioning is to start, keeping blade
vertical and at right angle to stem.
5. Keep sharp corner of cutting edge on slide with
cutting edge slanting upward toward you. This keeps
cutting edge sharp.
6. Move blade sideways against squared end of drop,
making thinnest section possible while watching
through dissecting microscope.
1. Put bryophyte in boiling water to relax it and select
several good, clean shoots.
2. Remove excess water by pressing the bryophyte
gently between absorbent paper towels or blotters.
3. Roll a 5-10 x 30-40 mm strips of hard-surfaced, thin
transparent tracing paper (such as that used by
architects) lengthwise into a tight scroll. The size
depends on the size of the strips. Open the roll and
place the bryophyte shoots longitudinally into the first
coil of the roll, using fine curved forceps.
4. Carefully roll the shoots up in the strip, using thumbs
and index fingers of both hands.
5. Hold this roll up to the light to locate the upper ends
of the shoots and grasp the roll just above the shoot
tip with a pair of forceps.
6. Lay a strip of good quality, smooth, white cardboard
(10x40 mm) parallel with the proximal edge of the
cutting block.
7. Place the bryophyte roll longitudinally upon the white
paper near the proximal edge, holding it down with
the left index fingernail at the shoot apex.
8. Using a sharp safety razor blade, cut off the anterior
portion of the paper roll and discard.
9. Begin cutting sections of stems and leaves through the
tracing paper, using your fingernail as a guide. After
each cut, move the blade back slightly before making
the next cut.
10. As sections are cut, dip the razor blade in a drop of
water on a glass slide to remove the sections.
11. Remove the sections of tracing paper from among the
leaves, adding a few drops of water to facilitate the
removal.
12. Excess water can be removed by holding the slide
over an alcohol lamp, leaving only a thin layer.
13. Large leaves like those of Polytrichum can be treated
in the same way as the stems.
Pith Sandwich Cutting Tool (Trotter 1955)
1. Cut a piece of pith from common elder (Sambucus
niger) 3-4 cm long x 1 cm wide. Make sure ends are
cut clean to make a cylinder.
2. Cut cleanly as possible with sharp safety razor blade
down the middle to avoid fraying.
3. Put drop of water on clean slide.
4. Lay half of pith on convex side.
5. Place dry specimen at end on flat side, slightly
extended beyond pith.
6. Make a sandwich by placing other half of pith flat
side onto the first flat side of pith, being careful to
align edges.
7. Hold sandwich firmly and dip end with moss into
water.
8. Place sandwich onto a glass slide without losing grip
and make a first cut close to end that holds moss,
using sharp, clean safety razor blade, and discard that
cut.
9. Dip to wet end of sandwich again.
10. Press firmly down on the pith above the specimen and
cut first section as thinly as possible next to the end
of the pith, taking care not to cut the pith. You may
want to do this while watching through a dissecting
microscope.
11. After making several cuts, use razor blade or
dissecting needle to move cut sections to opposite end
of slide and into drop of water or wetting agent.
12. Repeat until you have enough sections.
13. Place coverslip onto cut sections and water.
14. Add extra water at edge of coverslip if needed.
15. To examine, close down the diaphragm that controls
the light and examine on low power (40X total).
Note that sections that are visible to the naked eye are
probably too thick to be useful. Note also that stems with
oblique leaves, like Fissidens, may have to be placed with
stems in an oblique position so that leaves are
perpendicular to the cutting edge. Furthermore, plants with
very brittle cells, like Rhabdoweisia, may make it difficult
to get good sections.
Chopping Method
Some bryologists subscribe to a chopping technique.
They use a moist, but not flooded, stem with leaves, placed
on a glass slide. These are chopped with a razor blade from
the apex towards the base. Using some very fine forceps,
usually adding a very small amount of water (in addition to
that held between the tips by capillary action), they are
spread about the water drop where the coverslip will go.
After the coverslip is added, this preparation can be cleared,
if necessary, by infiltrating it with a drop or two of lactic
acid, and warming as discussed under Clearing Spores
above. I (Glime) have always felt this chopping method
was a waste of time since any chopped bits must be
examined afterwards, and often none of them is useful.
Most, if not all, of the sections will be wedge-shaped and
won't lie on their sides. Perhaps I just gave up too soon
before I perfected my skill.
Roll and Chop (Wilson 1990; Zander, Bryonet
8 July 2008)
Wilson (1990) presented a method he calls the "roll
and chop" method (Figure 53). He uses a dissecting needle
to hold the leaf or stem on a glass slide. After each cut, the
needle is rolled back a tiny bit and cut again with the razor
blade against the needle. I haven't tried this method, but I
do have a concern. If one starts cutting from the bottom of
the stem, the leaves become detached after the first cut,
2-2-21
reducing the chances they will subsequently be cut in thin
sections. If one starts at the tip, rolling the needle will butt
into leaf tips and roll under them instead of on top of them.
I asked Richard Zander for his advice on this, and he
agreed that if you start at the bottom of the plant the leaves
fall off. Rather, he always does "one leaf at a time if
possible, since results are better. Hold the leaf down, apex
away, then chop across the middle of the leaf while rolling.
A substitute for rolling the needle (probe) is to hold the leaf
down at an angle and slowly chop while dragging the blade
down the needle; results are the same. Sometimes one can
hold the whole plant down with a needle across the plant
apex at an angle perpendicular to the leaves and chop
across many leaves. This results in a mess but sometimes
cross sections result. Less tedious than doing one leaf at a
time, though."
Figure 53.
The roll-and-chop method of sectioning
bryophytes. This would usually be done while looking through a
dissecting microscope. Modified from Wilson (1990).
Richard Zander (Bryonet 8 July 2008) recommends
that single-edge razor blades (Figure 54) for sectioning
should be discarded after five to ten uses because they
become dull. He described his technique, essentially that
of Wilson, on Bryonet: "One holds a leaf or stem
crosswise with a stiff dissecting needle, then slices the
material with a razor blade held longitudinally against the
far side of the needle, meanwhile rolling the needle slowly
towards oneself to gradually expose uncut portions of the
material.
Figure 54.
Micromark.
Box of single-edge razor blades.
Photo by
2-2-22
Practice (and a relatively fresh blade) makes this
technique quite effective, even for very small leaves.
Remember to scrape off sections (especially stem sections)
adhering to the razor blade with a dissecting needle after
cutting. The usual pair of compound and dissecting
microscopes are needed, but using an additional illuminator
with the dissecting microscope for fine dissections rather
than just a single lamp will prove surprisingly
advantageous for observation of fine features." Zander and
others (Bryonet 8 July 2008) suggest Micromark
<http://www.micromark.com/> as a source for razor blades.
Modified Roll and Chop (Kellman 2005)
Kellman (2005) criticized this roll and chop method
because it is difficult and often produces sections that are
too thick. The pressure needs to be even and sufficient to
prevent the leaf from tearing. He recommended solving the
first problem of thick sections by making a special needle
using a sewing needle. The needle is cut to the desired
length (about 7.5 cm) and inserted into a 4 cm piece of a 1
cm wooden dowel by drilling a 1.6 mm hole into the end to
a depth of about 1.2 cm. The large diameter of the dowel
provides one with a better grip and makes it easier to roll
the needle a shorter distance. Kellman finds that the best
needle is a 7.5 cm (3") soft sculpture doll needle 1 mm in
diameter (Dritz product #56D). The cut end of the needle
should be dipped into a drop of glue and put into the hole in
the dowel. The short end of the needle can then be wedged
into the hole beside the needle to position it firmly.
Kellman warns that cutting the needle often results in
having the cut off end flying across the room, so he
recommends that it be cut inside a cloth or plastic bag so
that it can be retrieved easily. The next step creates the
tread that helps the needle grip the leaf. Run an emery
board or sandpaper along the length of the needle, rotate
the needle and repeat until the entire needle has a tread. Do
not run the emery board or paper around the needle because
that will not create the lengthwise treads needed.
Kellman solves the tearing and uneven pressure
problem by stacking several leaves on top of each other to
cut them. This also provides more sections, saving time.
1. To prepare the sections, place the stem on right-hand
side of a clean slide and remove leaves under a
dissecting microscope.
2. Select the leaves you want to section and move them
to the left side of the slide without adding more water.
3. Once you have moved the chosen leaves, stack them
together like spoons, stacking at least 3 leaves.
4. When the stack is ready, place the needle over the
stack, pressing down lightly.
5. Use a sharp blade to cut along the away side of the
needle. Use a chopping type of cut instead of a slice,
a method not feasible with a single leaf. The full edge
of the blade should reach the slide at one time.
6. Move the cut piece away and roll the needle as little
as possible back toward you.
7. Make another cut, making the first section.
8. If sections get stuck to the blade, place a drop of
water in the middle and dip the blade in it to remove
the sections.
9. Continue this procedure until you have enough
sections. Then make a slide as usual. You can place
a coverslip on the stem and remaining leaves on the
right to view whole leaves and another on the sections,
all on one slide.
Dissecting Microscope Hand Sections (Welch
1957; Schofield 1985)
This method works well for leafy stems, branches, and
large leaves. Some bryophytes, like Polytrichum, require
leaf sectioning to view special structures like the lamellae.
Because this is a large leaf, it is a good representative for a
beginner to use for practice. Welch (1957; Schofield
(1985) published the technique that works best for me
(Glime):
1. Place a wet Polytrichum leaf or leafy branch/stem on
a dry slide.
2. Put a drop of water on one side of the slide, away
from the leaf.
3. Cut away the tip with a sharp razor blade about 1/3
from the tip end of the leaf.
4. Discard your first cut.
5. While viewing through a dissecting microscope, cut
as close to the previous cut as possible. Use one hand
to cut and the other to guide and steady the cutting
hand while holding the specimen with a fingernail or
a pair of curved microforceps.
6. Cut 8-10 very thin sections and dip your razor blade
in the drop of water to free them.
7. Examine the sections with the dissecting microscope
to see if any of them are lying in cross section.
8. Continue cutting until you have about 30-40 sections.
9. If there are satisfactory sections, put a coverslip on
the slide and examine the leaves under low and high
power on the compound microscope.
With this technique I can usually get 5-8 sections that will
lie on their sides as they should.
Double Slide Sectioning Technique
Sean Edwards (pers. comm. 20 July 2012) has
provided us with his double slide sectioning technique,
based on his thesis (Edwards 1976 – See Edwards 2012).
The following description is only slightly modified from
his description.
This method allows, with very little practice, good
clean sections of about 10 µm thickness to be taken from
any required part of the moss leaf, with no preparation or
specialized equipment, and within a matter of seconds. The
equipment required is the normal laboratory dissecting
microscope (or good close eyesight), two 7.5 × 2.5 mm
standard glass slides, and a supply of double-edged or
single-edged razor blades.
Selected moist leaves are arranged parallel with each
other on a glass slide, with the parts to be sectioned aligned
as shown by the arrows in Figure 55. The second slide is
laid (with care) over the leaves, so that its long edge is also
aligned with the parts to be sectioned (Figure 56-Figure
57). This may be checked with a dissecting microscope if
necessary, and individual leaves adjusted. Firm pressure is
applied to the upper slide by the finger of one hand, and
half a double-edged razor blade is drawn with the other
hand across the leaves, using the upper slide as a guide
(Figure 56, Figure 57). Only a corner of the blade is used,
but if the 'angle of elevation' of the blade is sufficiently
small (about 15°-20°, perhaps less than that indicated in
Figure 56), the cut is perfectly clean.
2-2-23
even finer control over the thickness of the last few
sections. Although the sections must in theory be slightly
wedge-shaped because of 15° angle, this is not noticeable
in practice.
Figure 55. Placing specimen on slide in first step of the
double slide sectioning technique of Sean Edwards. Drawing by
Sean Edwards.
Figure 56. Alignment of slide and specimens in double slide
sectioning technique of Sean Edwards. Drawing by Sean
Edwards.
Figure 58. Cutting position of the razor blade in the double
slide sectioning technique of Sean Edwards. θ is the angle of tilt
and P1 is the fulcrum. P2 is the position of the fulcrum after the tilt
of the blade has passed 0° (vertical). Drawing by Sean Edwards.
Pressure on the blade has to be judged by experience,
but it should be no more than is necessary to cut the leaves.
One blade corner may provide many series of sections, but
such economy is usually not necessary; only with very old
and fragile material should a fresh corner be used for each
operation. It seems that an 'angle of elevation' of about
15°-20° enables the pressure to be taken by the less
vulnerable curved corner of the blade, while allowing the
razor edge unimpeded access to the leaves. It is clearly
advantageous to keep this angle constant. If the broken
corner of a half-blade immediately above the cutting corner
is bent somewhat, just before it is first used, then the
unused cutting corners can be recognized without
confusion, and a packet of ten blades can be used to section
at least forty plants. Particular advantages of this method
lie in the degree of control and inspection allowed before
and during cutting, by the transparency of the glass cuttingguide, and also in the world-wide availability and
cheapness of double-edged razor blades.
Figure 57. Sectioning setup of double slide sectioning
technique of Sean Edwards. Photo by Sean Edwards.
Sections are made by adjusting the tilt of the razor
blade for each successive cut; the first cut is made with the
blade leaning somewhat (about 15°) towards the upper
slide, and this angle is progressively lessened. The
situation is shown diagrammatically in Figure 58, where θ
is the angle of tilt and P1 is the fulcrum. The angle of tilt is
surprisingly easy to control, and even a relatively coarse
adjustment will give a fine control over the section
thickness. After the tilt of the blade has passed 0°
(vertical), the fulcrum moves down to P2, resulting in an
Figure 59. Cutting sections along edge of top slide in double
slide sectioning technique of Sean Edwards. Note cut sections in
water on the lower slide. Photo by Sean Edwards.
2-2-24
might try adding a bit of wetting solution inside the capsule
before sectioning. Be careful that the wetting agent does
not get on the ice mount because the water drops will run
off before they can freeze. (We haven't really tried this, so
we don't know if it will work.)
Figure 60. Cut sections along edge of top slide in double
slide sectioning technique of Sean Edwards. Note the alignment
of multiple stem pieces under top slide. Photo by Sean Edwards.
Cryostat Sections
If you are fortunate enough to have a cryostat, you can
get excellent, consistent sections. I inherited one that had
been obtained as government surplus. The principle is that
it freezes your specimen in ice. The specimen is prepared
by turning the cold stage to a very cool temperature and
building up an ice base with a few drops of water, waiting
for each drop to freeze before adding the next. Then the
specimen is placed there vertically. If you are cutting small
leaves, you may want to position several of them on the ice.
Once the specimen is positioned, continue to add drops of
water, letting each freeze before adding the next. Once you
have covered the portion of the specimen you need, you
can cut off any excess with a sharp razor blade. The disk is
then clamped into a holder in front of a blade. This blade
(or perhaps the holder) can be moved by "winding" much
like an old Victrola. Each time the blade comes down, it
cuts a narrow slice from the ice and bryophyte. These must
be collected on a cold, dry slide placed under the ice ribbon
created – something that must be done quickly. A pair of
microforceps can help to remove all the ribbon from the
blade. To make the slide cold, keep it inside the cryostat
while you are building the ice mound and doing the
sectioning. A warm slide will melt the ribbon immediately
and you can lose your slices.
The icy ribbon can be moved to the center of the slide
if done quickly before it melts. Then you can add a drop of
water and coverslip as you would for any slide.
The cryostat can be adjusted for the thickness of the
sections. The necessary thickness depends on the thickness
of the specimen (leaf, stem). Capsules are a bit more
difficult once they form an internal air chamber because the
air will be trapped inside. If this becomes a problem, you
Figure 61. Polytrichum juniperinum leaf section using a
cryostat. Photo by John Hribljan.
Stems and Small Leaves
Mosses lack lignified vascular tissue in their stems, but
they may have vascular elements called hydroids (waterconducting elements) and leptoids (photosynthateconducting elements). Additionally, the center of the stem
may contain small, thick-walled cells that serve as
strengthening tissue (Figure 62), but that does not seem to
have any conduction function. None of these structures can
be seen without sectioning the stem. Furthermore, it is
difficult to section small leaves by themselves, so they are
best sectioned on an intact stem or branch. This is the
method that works for me (Glime):
1. Place a wet moss stem on a dry slide.
2. Put a drop of water on one side of the slide, away
from the stem.
3. While viewing through a dissecting microscope, use a
sharp razor blade to cut as close to the end of the
stem as possible. Use a fingernail or finger of one
hand to guide (the one holding the stem) and steady
the hand holding the blade. Alternatively, you might
find it easier to press down on the stem with a pair of
curved forceps instead of holding it with your finger.
4. Discard your first cut.
5. Cut 8-10 very thin sections and dip your razor blade
in the drop of water to free them.
6. Examine the sections with the dissecting microscope
to see if any of them are lying in cross section,
indicating they are thin enough.
7. If there are satisfactory sections, put a coverslip on
the slide and examine the stems under low and high
power on the compound microscope.
8. Use a microscope with plane polarized light to see
cells with phenolic compounds in the stem.
Figure 62. Stem cross section of the moss Molendoa
sendtneriana showing central strand.
Photo by Dale A.
Zimmerman Herbarium, Western New Mexico University.
Techniques for Special Structures
Clearing Spores
Tom Blockeel (Bryonet 24 January 2012) sought a
method to make it easier to see the very dark or blackish
spores of species like those of Riccia (Figure 63). The
ornamentation of the spores can help in identification, but it
is not possible to observe it clearly with transmitted light.
Wagner (Bryonet 24 January 2012) suggested using a
combination of transmitted and reflected light. The
reflected light can be a strong LED light from a bicycle
headlamp. This, combined with stacked images, can
provide excellent quality (Figure 64).
Figure 63. Spore of Riccia sorocarpa showing its dark color
and density, preventing one from seeing spore wall details without
special techniques. Photo from EOL through Creative Commons.
2-2-25
Figure 64. Spore of Riccia sorocarpa using both transmitted
and reflected light plus stacking. Compare the clarity to that of
the same species in Figure 63. Photo by David Wagner.
Marko Sabovljevic (Bryonet 24 January 2012)
suggested using 5-10% bleach (NaOCl – 8% of active
chlorine) for 1-3 minutes to clear the spores, a method also
suggested by Richard Zander and Jörn Hentschel in the
same Bryonet thread. Hentschel also suggested calcium
hypochlorite (Ca(ClO)2), the C-Solution used by
lichenologists for their spot test. Martin Godfrey (Bryonet
25 January 2012) uses gum chloral to clear dark, dense
specimens and also make a permanent preparation. But
Upton (1993) reports that gum-chloral slides deteriorate
steadily with time and specimens become irretrievably lost.
Several bryologists (Richard Zander, Rod Seppelt, Bryonet
24 January 2012) also suggested lactic acid, but it wasn't
clear that they had actually tried it for black spores.
Seppelt also suggested a strong detergent like Tween 80
because it reduces the black pigment in some lichens. Tom
Blockeel reported that the bleach "does the trick perfectly
well!" (Bryonet 6 February 2012).
Rod Seppelt (Bryonet 14 November 1997) suggested
staining spores with malachite green, acid fuchsin, and
orange G, a method used for testing pollen (Alexander
1969). The viable pollen stains deep red-purple, whereas
the aborted pollen stains green. This recipe uses chloral
hydrate, a controlled substance in the US. The solution
uses 10 ml ethanol; 1 ml 1% malachite green in 95%
ethanol; 50 ml distilled water; 25 ml glycerol; 5 gm phenol;
5 gm chloral hydrate; 5 ml 1% acid fuchsin in water; 0.5
ml 1% orange G in water; and 1-4 ml glacial acetic acid
(for very thin to very thick walls). This should work as
well for spores.
SEM
Scanning Electron Microscopy (SEM) can reveal
details not visible with an ordinary light microscope.
Miyoshi (1969) demonstrated the intricate detail of
Schistostega pennata and Hedwigia ciliata by using the
Scanning Electron Microscope (SEM), compared to images
using the light microscope. The image in Figure 65 was
taken using SEM photography and can be compared to that
of the same species using ordinary light (Figure 63) or both
transmitted and reflected light (Figure 64). The SEM
technique is somewhat complex and time-consuming and
2-2-26
will not be covered at this time. Methods can be found in
Hofmann et al. 1996, Zhang et al. 2007, and Srivastava et
al. 2011, and many others.
Figure 65. Riccia sorocarpa distal spore wall SEM. Photo
by William T. Doyle.
Vacuoles
Many bryologists seemed to consider that bryophytes
did not have vacuoles, but it appears this is only true for
some taxa (Rod Seppelt, Bryonet 14 November 1997; Jeff
Bates, Bryonet 14 November 1997). It is interesting that
Seppelt reports that vacuoles seem to be absent in most
Antarctic mosses. This suggests that absence of vacuoles
may be an adaptation to cold temperatures. This would be
an interesting correlation to examine.
One indicator that a cell has a vacuole is the position
of the chloroplasts (Michael Christianson, Bryonet 14
November 1997). If they are crowded around the periphery
of the cell, it is likely that a vacuole is occupying the center
of the cell.
In Figure 66, fluorescent microscopy
demonstrates the position of the chloroplasts at the
periphery of the leaf cells of Funaria hygrometrica.
Figure 66.
Leaf of Funaria hygrometrica showing
chlorophyll fluorescence and demonstrating the clustering of
chloroplasts at the cell margins. Such positioning indicates the
presence of a vacuole. Photo by Janice Glime.
Liverworts
Oil bodies can be a problem because they disappear as
the liverwort dries (Tom Thekathyil, pers comm. 27 August
2012), in some species disappearing within hours despite a
moist state of hydration. David Wagner (Bryonet 5
September 2012) considers it a general rule that when cells
with oil bodies die, the oil bodies dissipate. Liverworts on
rotting logs (which are moisture sinks) never dry out in
nature, but when they dry, they die. Unlike other
bryophytes, they are not desiccation tolerant. Calypogeia
species must be examined for oil-body characters before
they dry. Once dry, the oil bodies are gone forever and
slow drying doesn't help. On the other hand, liverworts
that grow in extreme environments, like Marsupella spp.
on rocks in alpine situations, are as desiccation tolerant as
any bryophyte. If air dried, herbarium specimens will
retain oil bodies for years because the cells are NOT dead.
They live for years in a desiccated condition. To have any
chance of seeing oil-bodies in dried specimens, they must
be rehydrated slowly with plain water. Sometimes Wagner
has been surprised at getting good results. Also to be
remembered is that oil bodies can change character as they
age after collecting. There's no substitute for immediate
observation upon collections. There are some mysterious
anomalies. Scapania gymnostomophila has oil bodies that
persist for decades, itself a distinctive taxonomic character.
Nevertheless, liverworts survive wetting and drying in
nature. Rod Seppelt (Bryonet 27 August 2012) reported
that Jeff Duckett told him that the liverworts must dry
SLOWLY for the oil bodies to survive, but does this
always work, or does the death of oil bodies explain why so
many liverworts seem to require a moist environment?
Oil bodies are often essential for identification.
Several methods of liverwort preservation have been
suggested (Lehman & Schulz 1982; Ohta 1991). Lehmann
and Schulz suggest a method of fixation that preserves the
oil bodies, as do Müller-Stoll and Ahrens (1990). The
latter researchers provide a method of staining oil bodies in
live cells with diachromes and fluorochromes. If you can
read the language, these may be helpful.
Peristome Teeth
To study peristomes in plane polarized light, the
ventral and dorsal laminae of the teeth (not outer and inner
peristomes) must be separated (Taylor 1959). Examination
may even require viewing a cross section of a tooth.
1. Split capsule vertically with a razor blade.
2. Soak teeth in groups of 3-4 in 5% solution of
pectinase for 24 hours.
3. Wash in 3 or more baths of distilled water.
4. Make gum syrup mountant
A: 40 g gum arabic
0.5 g phenol crystals
60 cc water
B: 52 parts cane sugar
30 parts water (by volume)
Combine 25 cc A, 15 cc B, and 2 g glycerin.
5. Cover a small area of a slide with a thin coating of the
gum syrup mountant.
6. For peristomes, permit gum arabic to become almost
dry.
7. Transfer teeth in groups of 3-4 to mountant, making
sure some groups show the ventral and others dorsal
surface.
8. If peristome teeth curl, they can be moistened slightly
with a damp (not wet) fine water color brush (#
00000).
9. The gum arabic can be remoistened if needed,
especially if used for leaves and other structures.
10. The teeth can be flattened on the slide with a needle
or the damp brush.
11. Make sure the gum syrup is nearly hard, but soft
enough to flow under pressure. This will take
practice to prevent ripples from too much liquid, but
must permit the teeth to pull apart.
12. To make the teeth very flat (desirable), cover a part of
the slide lightly with a light coating of paraffin wax or
other substance to prevent the adhesive from sticking
to it.
13. Press the coated slide against the teeth until they are
tightly pressed against the mountant.
14. Permit the gum arabic mountant to harden.
15. Remove uppermost surface of lamina on each set of
teeth by gentle scraping, using a dull tool such as a
discarded side-cutting dental tool.
16. Remove the loosened particles with a dry brush.
17. Lightly moisten the gum syrup to get a smooth
surface and allow to dry.
18. To make the slide permanent, add the desired finisher,
such as gum-chloral.
2-2-27
Stains permit further clarification of structures such
as pores and wall markings and permit determination of
cell types. They can be as simple as food coloring or an
array of chemical stains used singly or in combination.
Identification of Sphagnum usually requires a stain to
discern the leaf cell pores.
Archegonia and
spermatogenous cells can be stained with fast green.
Fluorescent dyes coupled with a fluorescence
microscope can reveal bulbils and determine if spores
are viable. A pectinase preparation can be used to stain
liverwort capsules.
Some bryophytes (esp. Pottiaceae) produce
different colors in reaction to a mix of HCl, KOH,
concentrated H2NO3, and H2SO4. Some Sphagnum
subgenera respond to pH and have distinctive colors in
NaHCO3.
Removing leaves from stems is aided by a
dissecting microscope and microforceps. Sharp razor
blades can be used to make sections of leaves and
stems.
Cutting is best done under a dissecting
microscope, with the method being largely a matter of
preference, including chopping, wax mounts, pith
sandwich, cutting block, and double slide sectioning. If
you are lucky enough to have a cryostat, you can it to
make sections.
Some structures require special treatment, such as
clearing spores, using SEM, seeing vacuoles, preserving
and seeing oil bodies, and seeing details of peristome
teeth.
To View Teeth:
19. Place the finished slide on the rotating stage of a
polarizing microscope and turn stage to a position
where light is extinguished when viewing slide.
20. Insert gypsum tube into microscope tube and rotate
stage clockwise.
21. If tooth lamina becomes blue or green, chains run N-S
when tooth is returned to this extinction position.
22. If tooth lamina becomes yellow after rotation, search
for a position at right angles and repeat the test.
23. Be careful not to rotate counter-clockwise.
24. If all chains are parallel, you will not find the bright
color change, but usually at least some will show an
acute angle between two sets of chains.
Summary
Bryophytes often need to be cleaned before they
are mounted for observation. Methods for doing this
include a special bryophyte washing machine, netting
on an embroidery hoop, wash bottle, HCl, H2O2, and
agitation. Dried bryophytes need to be rehydrated using
a wetting agent such as water, soap, detergent, heated
water, 2% KOH, Pohlstoffe (docusate sodium), or
Agral 600. Some leaves need to be cleared before cell
wall papillae and wall structure can be seen clearly,
using reagents such as lactic acid, KOH, NaOH, or
chloral hydrate. Some species require air drying or
dehydration in ETOH to prepare them for making a
slide.
Acknowledgments
Numerous discussions by Bryonetters have contributed
heavily to this chapter. I appreciate the additional help
from Richard Zander who answered my many questions
quickly and thoroughly and alerted me to his websites.
Isawo Kawai kindly sent me numerous pictures and
information on his staining procedure. Ralf Wagner
provided me with the instructions for the Etzold staining
and W3A staining.
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2-2-30

Laboratory Techniques: Slide Preparation and

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